The electrical behavior of cells and cell membranes is of profound importance in basic research as well as in modern drug development. A specific area of interest in this field is in the study of ion channels and transporters [1]. Ion channels are protein-based pores found in the cell membrane that are responsible for maintaining the electrochemical gradients between the extracellular environment and the cell cytoplasm. These channels quite often are selectively permeable to a particular type of ion, e.g., calcium, chloride, potassium, or sodium. The channels generally comprise two parts: (1) the pore itself, and (2) a switch mechanism that regulates the conductance of the pore. The switch mechanism may be controlled by transmembrane voltage changes, covalent modification, mechanical stimulation, and/or chemical ligands (e.g., through the activation or deactivation of an associated membrane receptor), among others. Ion channels are passive elements in that, once opened, ions flow in the direction of existing electrochemical gradients. Ion transporters are similar to ion channels in that they are involved in the transport of ions across cell membranes; however, they differ from ion channels in that they require energy for their function and in that they tend to pump actively against established electrochemical gradients.
Ion channels are prevalent in the body and are necessary for many physiological functions, including the beating of the heart, the contraction of voluntary muscles, and the signaling of neurons. They also are found in the linings of blood vessels, allowing for physiological regulation of blood pressure, and in the pancreas, allowing for the control of insulin release. As such, the study of ion channels is a very diverse and prolific area encompassing basic academic research as well as biotechnical and pharmaceutical research. Experiments on ion channels may be performed on cell lines that endogenously express the ion channel of interest (“native channels”) as well as on recombinant expression systems such as the Xenopus oocyte or mammalian cell lines (e.g., CHO, HEK, etc.) that have been transiently or stably transfected to express the ion channel by well-known techniques [2, 3]. Electrophysiology also is performed on isolated cell membranes or vesicles as well as on synthetic membranes where solubilized channels are reconstituted into a manufactured membrane [4].
I. Instrumentation
To date, the most useful and widely utilized tool for the study of ion channels and transporters is a technique called “patch clamping.” This technique was first introduced almost 25 years ago [5-7], and consists of using a small glass capillary to function as an electrode in measuring currents and voltages from individual cells. FIG. 1 shows a typical patch clamp measurement geometry. A glass capillary 2 is first heated and pulled to a fine tip. The capillary is then filled with a saline buffer solution 4 and fitted with a Ag/AgCl electrode 6. The function of the Ag/AgCl electrode is to provide an electrical connection to a wire via the reversible exchange of chloride ions in the pipette solution.
Through the use of a microscope and micromanipulating arm (not shown), the user finds a biological cell or cell membrane 8 containing ion channels 10 of interest and gently touches the cell membrane with the pipette. The measurement circuit is completed via the external ionic solution 12 and a second Ag/AgCl bath electrode 14. A high-impedance operational amplifier 16 senses the current flowing in the circuit, which is subsequently recorded and analyzed with a data recording system 18. A key to the successful function of the technique is the ability to form a high electrical resistance (˜1 GΩ) seal between the glass pipette and the cell membrane 20, so that the current recorded by the amplifier is dominated by ions 22 flowing through the cell membrane and not by ions flowing around the glass pipette directly into the bath solution.
Once a high-resistance seal is achieved between the pipette and the cell membrane, there are many measurement configurations that the system can take, including the “whole-cell,” “perforated-patch,” and “inside-out” patch clamp configurations. The whole-cell voltage clamp is one of the more common configurations. In the whole-cell voltage clamp, the portion of membrane at the end of the pipette 24 is permeabilized so as effectively to place the pipette electrode inside the cell. This, in turn, allows for an external voltage command 26 to be placed between the intracellular pipette electrode and the extracellular bath electrode, thereby providing control of the cell's transmembrane voltage potential. The term “whole cell” is derived from the fact that, with this configuration, the instrument measures the majority of the currents in the entire cell membrane.
The electrical permeabilization of the membrane at the end of the pipette can be induced in many ways. Permeabilization often is achieved by using voltage pulses of sufficient strength and duration that the membrane inside the pipette physically breaks down. This approach is well known in the field and is commonly referred to as “zapping” [8]. Permeabilization also may be achieved by using certain antibiotics, such as Nystatin and Amphotericin B [9]. These antibiotics work by forming chemical pores in the cell membrane that are permeable to monovalent ions, such as chloride. Since chloride is the current-carrying ion for the commonly used Ag/AgCl electrode, these antibiotics can produce a low resistance electrical access to the interior of the cell. The advantage of the chemical technique is that the membrane patch remains intact so that larger intracellular molecules remain inside the cell, rather than being flushed out by the pipette solution as with the zapping technique. This approach also is well known in the field and is commonly referred to as a “perforated patch” [8-10].
The formation of high-resistance electrical seals enables the measurement system to detect very small physiological membrane currents (e.g., ˜10−12 A). In addition, by perforating a portion of the cell membrane either electrically or chemically, it is possible to control the voltage (voltage clamp) or current (current clamp) across the remaining intact portion of the cell membrane. This greatly enhances the utility of the technique for making physiological measurements of ion channel/transporter activity, since quite often this activity is dependent on transmembrane voltage. By being able to control the trans-membrane voltage (or current), it is possible to stimulate or deactivate ion channels or transporters with great precision and as such greatly enhance the ability to study complex drug interactions.
The development of the patch clamp technique revolutionized the field of electrophysiology, allowing for the direct electrical measurement of ion channel/transporter events in living cells, cell membranes, and artificial membranes. However, existing patch clamp techniques require operators with high levels of manual dexterity who must learn to record data from single cell or membrane preparations using a small glass capillary positioned under a microscope by a micromanipulating arm. Moreover, even skilled operators typically require tens of minutes to complete a single recording session, while, in the case of drug screening, it generally is preferable to obtain a new cell sample for each different chemical entity to be tested. Thus, existing techniques are not capable of looking at thousands of different conditions (e.g., chemical stimuli) per day, a common need in the biotechnical or pharmaceutical industry.
U.S. Pat. No. 6,063,260 to Olesen describes a system intended to improve the throughput and decrease the fluid volume required of standard patch clamp technology. The improvement relies on using a standard HPLC autosampler apparatus integrated into a standard patch clamp arrangement to more easily inject multiple fluids samples into the measurement system. The invention claims to increase throughput by making multiple sequential fluid additions to the same biological membrane faster and easier. However, the Olesen invention is deficient in several respects. First, it does not allow for a plurality of different biological samples to be measured simultaneously. Second, it does not eliminate the labor-intensive aspects of micromanipulation involved in standard patch clamp electrophysiology. Third, it does not address cases in biological drug screening where multiple chemical reagent additions to the same biological sample are to be avoided (as in the case of high-throughput drug screening).
Published PCT Application No. WO 99/66329 discusses the use of a perforated screen to conduct tests on biological materials, but the proposed system has significant, severe limitations in terms of practical implementation. For example, all embodiments discussed in the WO 99/66329 application utilize multiple apertures per fluid well, placing reliance on the growth of confluent cell matrices to effectuate sealing of the multiple perforations formed in relatively thick material. In addition, although the published application makes reference to automation, no workable, fully integrated systems are disclosed that are capable of high throughput and reliability.
The invention may address these and/or other shortcomings by providing instrumentation for automated, high-throughput studies of ion channels.
II. Ion Channel Assays
The rapid and diverse signaling kinetics of ion channels makes their study both interesting and technically challenging. Many ion channels can be activated and then deactivated in a few milliseconds. This rapid time scale implies that the instrumentation used to study channel kinetics should have a fairly high frequency bandwidth, for example, on the order of 10 kHz. Fortunately, such bandwidths are attainable, since high-bandwidth operational amplifiers are readily available. Unfortunately, this rapid time scale further implies that the method of stimulating ion channel events also should be fast.
The needed time scale of the stimulus depends in part on whether the channels are voltage gated or ligand gated. Voltage gated channels are activated or deactivated by changes in transmembrane voltage, as mentioned previously. For these channels, the same electronics used to record ion channel currents also can be used to control the voltage stimulus, since the time bandwidth of the stimulus, an electrical signal, is inherently fast enough to avoid degrading the kinetics of the voltage-gated ion channel signals. In contrast, ligand-gated channels are activated or deactivated by chemical or ligand binding. These channels may be gated by specific chemical messengers, such as the release of intracellular calcium, adenosine 3′,5′-monophosphate (cyclic AMP or cAMP), or acetylcholine (ACh), among others. In some cases, the chemical activation of an ion channel is extracellular in its initiation, and, in other cases, the chemical activation is intracellular. This implies that it is important that the compound not only can be released on the time scale of tens of milliseconds, but in some cases that the compound can be introduced within the membrane of a living cell.
The invention may address these and/or other shortcomings by providing channel assays for automated, high-throughput studies of voltage and/or ligand-gated ion channels.